Cellular Trafficking of Sn-2 Phosphatidylcholine Prodrugs

Precisionnanomedicine.com DOI: 10.29016/180730.1 @2018 Andover House, Andover, MA USA

The official Journal of CLINAM License: CC BY-NC-SA 4.0 128

Open Access Research Article

Prec. Nanomed. 2018 July;1(2):128-145.

Cellular Trafficking of Sn-2 Phosphatidylcholine Prodrugs Studied with Fluorescence Lifetime Imaging and Super-resolution Microscopy

Dolonchampa Maji PhDa,b *, Jin Lu PhDc, Pinaki Sarder Phdd, Anne H. Schmieder MSe, Grace Cui

MSe, Xiaoxia Yang BSe, Dipanjan Pan PhDf, Matthew D. Lew PhDc, Samuel Achilefu PhDa,b and

Gregory M. Lanza MD PhDb,e,†

aOptical Radiology Lab, Department of Radiology, Washington University School of Medicine, St. Louis, MO

63110, USA bDepartment of Biomedical Engineering, Washington University in St. Louis, MO 63130, USA

cDepartment of Electrical and Systems Engineering, Washington University in St. Louis, St. Louis, MO 63130,

USA dDepartment of Pathology and Anatomical Sciences, Jacobs School of Medicine & Biomedical Sciences,

University of Buffalo, Buffalo, NY 14203 eDivision of Cardiology, Department of Medicine, Washington University School of Medicine, St. Louis, MO,

USA fDepartment of Bioengineering, University of Illinois at Urbana-Champaign, Champaign, IL, USA

* Current address: Department of Neurological Surgery, Northwestern University, Feinberg School of

Medicine, Chicago, IL 60611

Submitted: July 18, 2018; Accepted: July 30, 2018; Posted August 3, 2018

Graphical abstract

Fluorescence lifetime imaging microscopy (FLIM) and single-molecule super-resolution microscopy (SRM) illustrate the

intracellular fate of Sn-2 phosphatidylcholine prodrugs.

† Corresponding author: Gregory M. Lanza, M.D. Ph.D. Division of Cardiology, Campus Box 8215,

660 Euclid Ave, Washington University School of Medicine, St. Louis, MO 63108 Tel: 314-454-8813,

Fax: 314-454-5265. Email: [email protected]

mailto:https://precisionnanomedicine.com/article/16

https://creativecommons.org/licenses/by-nc-sa/4.0/

mailto:[email protected]



Abstract

While the in vivo efficacy of Sn-2 phosphatidylcholine prodrugs incorporated into targeted, non-

pegylated lipid-encapsulated nanoparticles was demonstrated in prior preclinical studies, the

microscopic details of cell prodrug internalization and trafficking events are unknown. Classic

fluorescence microscopy, fluorescence lifetime imaging microscopy, and single-molecule super-

resolution microscopy were used to investigate the cellular handling of doxorubicin-prodrug and

AlexaFluor-488-prodrug. Sn-2 phosphatidylcholine prodrugs delivered by hemifusion of

nanoparticle and cell phospholipid membranes functioned as phosphatidylcholine mimics,

circumventing the challenges of endosome sequestration and release. Phosphatidylcholine prodrugs in

the outer cell membrane leaflet translocated to the inner membrane leaflet by ATP-dependent and ATP-

independent mechanisms and distributed broadly within the cytosolic membranes over the next 12 h.

A portion of the phosphatidylcholine prodrug populated vesicle membranes trafficked to the perinuclear

Golgi/ER region, where the drug was enzymatically liberated and activated. Native doxorubicin entered

the cells, passed rapidly to the nucleus, and bound to dsDNA, whereas DOX was first enzymatically

liberated from DOX-prodrug within the cytosol, particularly in the perinuclear region, before binding

nuclear dsDNA. Much of DOX-prodrug was initially retained within intracellular membranes. In vitro anti-proliferation effectiveness of the two drug delivery approaches was equivalent at 48 h, suggesting

that residual intracellular DOX-prodrug may constitute a slow-release drug reservoir that enhances

effectiveness. We have demonstrated that Sn-2 phosphatidylcholine prodrugs function as

phosphatidylcholine mimics following reported pathways of phosphatidylcholine distribution and

metabolism. Drug complexed to the Sn-2 fatty acid is enzymatically liberated and reactivated over many

hours, which may enhance efficacy over time.

Keywords:

Phosphatidylcholine prodrugs, Nanomedicine, Fluorescence lifetime imaging microscopy, Super-

resolution microscopy

Abbreviations:

• AF488 AlexaFluor 488

• API active pharmaceutical ingredient

• αvβ3 alpha v beta 3 integrin

• CFDD Contact Facilitated Drug Delivery

• cPLA2 cytosolic phospholipase A2

• DAPI 4',6-Diamidino-2-phenylindole, dihydrochloride

• DMEM Dulbecco's Modified Eagle’s Medium

• dsDNA double-stranded deoxyribonucleic acid

• ER endoplasmic reticulum

• FLIM fluorescence lifetime imaging microscopy

• MC540 merocyanine 540

• PAINT Point Accumulation for Imaging in Nanoscale Topography

• Paz-PC 1-palmitoyl-2-azelaoyl-sn-glycero-3-phosphocholine"

• PC phosphatidylcholine

• PD prodrug

• PFOB perfluoro-1-bromooctane

• PE phosphatidylethanolamine

• PE-PTX phosphatidylethanolamine-paclitaxel

• PLA phospholipase

• SRM super-resolution microscopy

• Sn-2 stereospecific numbering position 2

• τamp amplitude average lifetime





Rationale and purpose

Sn-2 phosphatidylcholine prodrugs in non-

pegylated lipid nanoparticle membranes are

known to be resistant to premature release and

metabolism in circulation. This approach using

targeted drug delivery has been demonstrated to

be effective in a variety of preclinical models.

Cellular uptake of these compounds is

hypothesized to be secondary to a hemifusion

contact-facilitated drug delivery mechanism

with cytosolic enzymatic drug release, which

circumvents sequestration in the endosome-

lysosome pathway. This research utilized

advanced complementary optical imaging

methods, including confocal fluorescence

lifetime and super-resolution single molecule

microscopy, to elucidate and compare PC

prodrug cellular trafficking and metabolism

with previously known paths of natural PC.

Introduction

The concept of achieving Paul Erhlich’s

inspired vision of a “magic bullet” to treat

disease remains challenging in the context of

nanomedicine. While specific reasons vary

among nanosystems, in general, early systemic

drug loss, low net drug delivery to targeted

cells, and limited free drug intracellular

bioavailability have been major barriers [1].

Very hydrophobic drugs, such as fumagillin,

incorporated into the phosphatidylcholine-

based surfactant membranes of αvβ3-targeted

nanosized emulsion particles [2-7] provided

anti-angiogenic benefits that were broadly

demonstrated in vitro and in vivo by imaging,

microscopy, and functional outcomes [3, 6-8].

Hydrophobic drugs dissolved into the lipid

surfactant spontaneously transferred to targeted

cell membranes through an irreversible

hemifusion process referred to as contact-

facilitated drug delivery (CFDD) [2, 4]. Despite

in vivo effectiveness, the pharmacokinetics of

the drug relative to the other lipid surfactant components and the perfluorocarbon (PFC)

core were discrepant and showed marked

premature fumagillin loss during circulation

[9]. Lipid-anchored homing ligands and

gadolinium chelates co-incorporated into the

surfactant of PFC nanoparticles (NP) had

similar pharmacokinetic clearance rates as a

percentage of injected dose in rodents. Less

hydrophobic drugs dissolved into the lipid

surfactant, such as paclitaxel or doxorubicin,

had rapid circulatory clearance rates compared

to the PFC NP core and other membrane

constituents. The very poor particle retention of

these drugs accounted for their lack of efficacy

in cancer models when studied in vivo [10].

In an earlier report, rhodamine-PE was

employed as a fluorescent drug surrogate to

illustrate and elucidate the CFDD mechanism.

Rhodamine-PE delivered with integrin-targeted

PFC NP integrated into the cell membrane upon

binding and readily passed via lipid rafts into C-

32 human melanoma cells at 37C [4]. In

contradistinction to rhodamine-PE, paclitaxel

conjugated to phosphatidylethanolamine failed

to translocate from the outer to inner cell

membrane leaflets. The concept of chemically coupling drugs to the Sn-2 fatty acid of

phosphatidylcholine (PC) was envisioned as an

alternative approach to facilitate nanoparticle

drug retention during circulation, to protect

chemically-sensitive compounds from

interactions with blood constituents, and to

transfer into the targeted cell via CFDD. In vivo

studies demonstrated that the Sn-2 prodrug

motif was efficacious for fumagillin, docetaxel,

and recently cMYC-inhibitors [9-12].

While the concept of Sn-2 phospholipid

prodrugs was never considered for ligand-

targeted drug delivery in association with the

CFDD hemifusion mechanism, precedence for

Sn-2 phospholipid prodrugs was found to exist,

initially reported by David Thompson et al. in

the context of triggered drug release

mechanisms [13]. Later, Andresen and

colleagues [14-24] pursued an approach to

deliver chemotherapeutics as Sn-2 prodrugs via

untargeted liposomes, anticipating that

phospholipases liberated locally by cancers

would trigger local drug release. Their

hypothesis was essentially incorrect. Physical-

chemical modeling by this team revealed that

non-pegylated liposomes were resistant to free

water penetration and premature hydrolysis of

the synthetic ether-lipid prodrugs. However,

pegylation of liposomes, a design feature

commonly employed to extend circulatory half-

life, wicked water into the membrane creating

access to lipases with resultant rapid premature

drug release.

Phosphatidylcholine (PC) populates all cell

and intracellular membranes in contrast to phosphatidylethanolamine or





phosphatidylserine, which preferentially reside

in the inner cell membrane leaflet and other

specific organelles, such as the mitochondria

[25, 26]. The Sn-2 PC prodrug motif was

adopted as a PC mimic that would circumvent

endosomal NP internalization and resultant

losses by intercalation into the cell membrane

and trafficking throughout the intracellular

membranes until lipases, such as phospholipase

A2, liberated a bioactive drug. However, this

working hypothesis for the effectiveness of Sn-

2 PC prodrug has never been elucidated.

In the present study, complementary

microscopic imaging techniques were utilized

to explore cell membrane uptake, cellular

internalization and intracellular distribution of

a doxorubicin Sn-2 PC prodrug or

AlexaFluor™ Sn-2 PC prodrug. The objectives

were to compare the biopotency of a

phosphatidylcholine doxorubicin prodrug

(DOX-PD) with free doxorubicin (DOX) and to

utilize fluorescence lifetime imaging

microscopy (FLIM) to compare the trafficking

of DOX-PD versus free DOX from the outer

cell membrane to the nucleus. To corroborate

and extend the FLIM results, single-molecule

super-resolution optical microscopy [27-29]

was employed to map Sn-2 AlexaFluor 488

prodrug (AF488-PD) trajectories from the cell

membrane uptake, outer to inner membrane

leaflet transfer, and throughout the intracellular

membranes.

Experimental design

The present study employed traditional

fluorescence microscopy, confocal

fluorescence lifetime imaging microscopy

(FLIM), and single molecule super-resolution

optical microscopy in a complementary fashion

to track optically active Sn-2

phosphatidylcholine prodrug in normal and

cancerous cells. The experimental protocols

utilized a doxorubicin prodrug for fluorescence

microscopy and confocal fluorescence lifetime

imaging for assessments of the cell membrane,

cytosol, and nucleus. A more photostable dye,

AlexaFluor™ 488, was synthesized into an Sn-

2 prodrug motif to detect single prodrug

molecules in the presence of autofluorescence.

This enhanced photostability extended the

SRM tracking time and permitted better

estimates of molecular migration patterns and

prodrug membrane retention.

Fluorescence microscopy differentiated free

DOX and DOX-PD distribution from the outer

cell membrane, cytoplasm and nucleus. FLIM,

which assessed distributions between local

biochemical environments based on changes in

excited-state lifetimes, offered qualitative

discrimination between drug retained in cell

membranes, enzymatically released into the

cytosol, or in the nucleus bound to dsDNA.

Super-resolution optical microscopy

dynamically tracked single prodrug (AF488-

PD) molecules across multiple focal planes,

demonstrating spatial uptake, translocation and

distribution within the cell until the dye was

enzymatically liberated, after which rapid

molecular motion beyond the confocal volume precluded detection. Collectively, these

observations and measurements demonstrate

the CFDD drug delivery mechanism of PC

prodrugs and the avoidance of endosomal

sequestration and degradation.

Materials and methods

Synthesis and characterization of

doxorubicin and AlexaFluor 488 prodrugs

Doxorubicin Sn-2 prodrug conjugates

(DOX-PD) were synthesized in our laboratory

starting from commercially available

doxorubicin (Sigma Aldrich, St. Louis, MO)

and oxidized phospholipids (NOF America,

White Plains, NY). In this approach,

doxorubicin (0.22 mM) was dissolved in

dimethylsulfoxide (DMSO) and triethylamine

(1 mM). Added to this solution was (S)-2-(8-

carboxyoctanoyloxy)-3-(palmitoyloxy)propyl

2-(trimethylammonio) ethyl phosphate

(PAzPC, 0.2 mM, NOF America) dissolved in

chloroform with cyclohexylcarbodiimide (1

mM). The reagent mixture was shaken

overnight at room temperature (RT) then

purified using silica gel column

chromatography. The desired product DOX-PD

was characterized using mass spectroscopy:

Chemical Formula: C60H91N2O20P, ESI-MS

M+H+ Cal. 1190.63, exp.1191.6 (Shimadzu

LCMS 2010A, Kyoto, Japan) (Figure S1).

The AlexaFluor 488 Sn-2

phospholipid conjugate (AF488-PD) was

synthesized from commercially available

AlexaFluor 488 (Thermo Fisher Scientific,

Waltham, MA) and oxidized





phosphatidylcholine (NOF America, PAzPC).

In this approach, AlexaFluor 488 cadaverine

(1.56 µmoles in 1 mL dimethylformamide) was

combined with excess PAzPC (6.6 µmole)

dissolved in 2 mL chloroform to which

triethylamine (50 µl) and N,N’-diisopropylcarbodiimide (50 µL) were added.

The mixture was reacted for 2-3 hours.

Separation with silica TLC (0.2M

NH4Ac:MeOH:water - 20:100:200) revealed

only the conjugated dye (RF 0); the free dye

control was seen at RF ~ 0.8. The sample was

dialyzed (MWCO 500-1000) then lyophilized.

Purified AF488-PD was characterized using

mass spectroscopy: final chemical formula was

C74H113N7NaO23PS2+2H+, ESI-MS M+H+ Cal.

793, exp.793.85 (Shimadzu LCMS 2010A,

Kyoto, Japan) (Figure S1).

Synthesis and characterization of αvβ3 DOX-PD PFOB nanoparticles

Nanoparticles (NP) were prepared as a

microfluidized suspension of 20% (v/v)

perfluoro-1-bromooctane (PFOB, Exfluor

Research Corporation, TX), 2.0% (w/v) of a

surfactant co-mixture and 1.7% (w/v) glycerin

in Milli-Q® water. An αvβ3-integrin antagonist,

a quinolone nonpeptide developed by Bristol-

Myers Squibb Medical Imaging (US patent US

6511648 and related patents) was used for

homing (a gift, Kereos, Inc., St. Louis, MO).

The surfactant co-mixture included:

approximately 97.6 mole% lecithin, 0.15

mole% of αvβ3-ligand-conjugated lipid and 2.28

mole% of DOX-PD (0.5 mM). The surfactant

components were combined with the PFOB,

buffer and glycerine and Milli-Q® water, then

the mixture was homogenized at 20000 psi for

4 min. The NP were preserved under inert gas

in sterile sealed vials until use. Nominal αvβ3-

DOX-PD PFOB NP and size by dynamic light

scattering was 260 nm, with a polydispersity of

0.14 and zeta potential of -7.4 mV. The

surfactants of fluorescent PFOB NP

incorporated 0.1 mol% rhodamine-

phosphatidylethanolamine (Avanti Polar

Lipids, Alabaster, AL) or 0.1 mol% AF488-PD

in the lipid commixture at the expense of

phosphatidylcholine.

Absorption and fluorescence spectroscopy

DOX and DOX-PD were used for absorption

and fluorescence spectroscopy. The materials

were diluted in Milli-Q® water. Absorption

spectra were measured on a DU 640

spectrophotometer (Beckman-Coulter, Brea,

CA). Fluorescence emission spectra were

recorded on a FluoroLog 3 spectrofluorometer

(Horiba Jobin Yvon, Edison, NJ) using 585

nm/600-700 nm as excitation/emission

wavelength with 5 nm slits.

Cell culture

For fluorescence microscopy, 2F2B mouse

endothelial cells (ATCC CRL2168, Manassas,

VA, USA) were cultured in Dulbecco's

modified Eagle's medium (DMEM, no phenol

red, Gibco, Thermo Fisher Scientific, Waltham,

MA) with 4.5 g/L glucose and 10% heat

inactivated fetal bovine serum. Cells were

plated on sterile 12 mm cover glasses (#1.5) in

a 24 well plate at approximately 1x105 cells per

well and incubated overnight at 37 °C in a 5%

CO2 incubator. Doxorubicin (Sigma, St Louis,

MO, USA) or αvβ3-DOX-PD PFOB NPs were

added to the cells at a final concentration of 3

µM active drug in complete media. After 1 h,

the cells were washed three times in PBS and

fixed in 4% paraformaldehyde solution for 30

min. The coverslips were rinsed in PBS; the

cell nuclei were stained blue with DAPI (4',6-

diamidino-2-phenylindole, dihydrochloride);

and samples were analyzed by fluorescence

microscopy (Olympus BX61, Shinjuku, Tokyo,

Japan).

For confocal fluorescence lifetime imaging

microscopy, human C-32 melanoma cells

(Washington University Tissue Culture

Support Center, St. Louis, MO) were cultured

in Minimal Essential Media containing Earle’s

salts (no phenol red, Gibco, Thermo Fisher

Scientific, Waltham, MA) and supplemented

with 10% (v/v) fetal bovine serum (Sigma-

Aldrich, St. Louis, MO), 1% penicillin and 1%

streptomycin (Gibco). The cells were

trypsinized and plated on glass bottom dishes

(35 mm, glass #1, MatTek Corporation,

Ashland, MA) and grown overnight in phenol

red-free media. The drug formulations were

diluted in the culture media and cells were

incubated for varying times. After incubation

and washing, fluorescence lifetime imaging

was performed on live cells in a humidified

stage-top incubator with 5% CO2.

For super-resolution imaging, 2F2B mouse

endothelial cells were cultured in DMEM with





10% fetal bovine serum (Gibco) plus 1%

penicillin and 1% streptomycin (Gibco) and

incubated at 37ºC with 5% CO2. 2F2B cells

were trypsinized and plated on ozone-

pretreated high-tolerance coverslips (No. 1.5,

170 ± 5 μm thickness, Azer Scientific,

Morgantown, PA), and incubated for 48 h. The

cells were further treated with 1.6 μg/mL

AF488-PD at 37 ºC for 12 h, or 0.05 μg/mL

AF488-PD at room temperature for 5 min

before imaging. The 2F2B cells were also

treated with 1.4 μM PE-rhodamine at 37 ºC for

1 h and rinsed with PBS for standard

epifluorescence imaging.

Cytotoxicity assay

The retained cytotoxicity of DOX-PD

was compared to native DOX in mouse

endothelial cells. 2F2B cells were seeded on a

96 well plate (2000-5000 cells/well) in DMEM

with 10% heat-inactivated fetal bovine serum

plus 10 µM angiotensin II (Sigma-Aldrich, St.

Louis, MO) then incubated at 37 ºC with 5%

CO2. After 24 h, cells were incubated for 1 h

with a) αvβ3-DOX-PD PFOB NP (0.3 µM DOX

PD), b) equimolar equivalent of native

doxorubicin HCl or c) αvβ3-No drug NP. After

incubation the wells were washed three times

with PBS and returned to the incubator for 48

h. Cell proliferation was measured using the

MTT assay for cytotoxicity, which measures

the activity of cellular enzymes that reduce the

tetrazolium dye to the insoluble formazan

(Invitrogen; Thermo-Fisher Scientific,

Waltham, MA). Each treatment was replicated

6 times.

Fluorescence lifetime spectroscopy, imaging and data analysis

Confocal fluorescence lifetime spectroscopy

and imaging was performed with a MicroTime

200 microscope (Picoquant, Berlin, Germany).

For spectroscopy, fluorescence lifetimes of

DOX (50 µM) and DOX-PD (50 µM) solutions

in MilliQ® water (final DMSO 0.5%) were

measured under a 100X oil immersion

objective with 485 nm (15 au) laser excitation

pulsed at 40 MHz. Decay data were collected

through a 519 nm long pass filter and

accumulated until peak photon counts reached

10,000. Almost identical conditions were

applied for αvβ3-DOX-PD PFOB NPs (25 µM

doxorubicin concentration) except the pulse

rate was reduced to 20 MHz in order to capture

its slower decay. Live cell imaging utilized 485

nm laser excitation (~1000 au) at 20 MHz on

cells maintained in a stage top CO2 incubator

for the period of imaging.

Lifetime decay data collected from

spectroscopy and microscopic imaging were

analyzed on the SymphoTime software

(Picoquant, Berlin, Germany). All decays were

fit using multi-exponential tail fit using the

equation below:

𝑦(𝑡) = ∑ 𝐴[𝑖]exp {−𝑡

𝜏[𝑖]

𝑛−1

𝑖=0

} + 𝐵𝑘𝑔𝑟

Equation 1

Where: n = number of exponentials (n ≤ 2 for

all cases depending on goodness of fit); A =

amplitudes; 𝜏 = lifetimes; Bkgr = correction for

background (after-pulsing, dark counts,

environmental light).

Spectroscopy data were fit to mono- or bi-

exponential decay fits (depending on goodness

of fit). For cell images, raw decay data from all

cellular pixels were fit to bi-exponential models

and this method was used to generate the

corresponding lifetime images.

For cell compartment specific analyses,

nuclear, perinuclear and extra-perinuclear

regions of interest (ROIs) were drawn to

achieve 1000 or more counts to ensure effective

bi-exponential decay analysis. Bi-exponential

tail fit was used to fit decays from each ROI and

individual components (lifetime and amplitude)

were obtained. Good χ2 values (0.8-1.2) and

random distribution of x-axis residuals were

considered necessary for an accurate decay fit.

Amplitude average fluorescence lifetimes (τamp)

were calculated and reported using the

following model and reported as average

lifetime values.

𝜏𝑎𝑚𝑝 = ∑ 𝐴[𝑘]𝜏[𝑘]

𝑛−1

𝑘=0

∑ 𝐴[𝑘]

𝑛−1

𝑘=0

⁄

Equation 2

Analyzed data was plotted using Prism

(Graphpad, La Jolla, CA). Wherever required,

appropriate statistical analysis was performed





using Prism’s built-in functions and

corresponding p values were reported.

Super-resolution imaging and data analysis

Single-molecule super-resolution imaging

was used to image the binding and cellular

uptake of individual prodrug molecules. First,

images of individual fluorescent molecules

with switchable bright-dark states (blinking)

were captured, and their positions were

measured by fitting each blinking event to a

two-dimensional Gaussian function. A super-

resolved image was then reconstructed by

combining all positions from the stochastic

blinking events of fluorescent molecules. To

resolve the fluorescence signal from a single

molecule, a bright fluorophore, AlexaFluor

488 (AF488), was used and conjugated to the

phospholipid at the Sn-2 position as the prodrug

analog for super-resolution imaging. AF488-

PD was more photostable than DOX-PD and

permitted detection of single copies of AF488-

PD in the presence of autofluorescence,

extending SRM tracking time and improving

estimates of molecular migration patterns and

prodrug membrane retention. Freely-diffusing

AF488-PD in the extracellular buffer and in the

cytosol were not captured, since their quickly-

moving fluorescence signals were out of focus

and/or averaged out by the detector. However,

when AF488-PD temporarily docked onto the

lipid membrane, its slowed diffusion was

captured and localized from which a super-

resolution image was reconstructed with spatial

resolution beyond refraction limit. In this way,

single-molecule super-resolution imaging

precisely localized the prodrug molecules with

single-molecule sensitivity.

Super-resolution imaging was performed on a

home-built microscope equipped with a 100X

oil immersion objective (Figure S2) [30, 31].

After AF488-PD incubation, the cells were

washed with PBS to remove the excess prodrug

in the media. Cells were first illuminated with a

488-nm laser (OBIS, Coherent) at a peak

intensity of 0.12 kW∕cm2 at the sample. The

images were filtered with a 523/610 dual-band

bandpass filter (Semrock FF01-523/610) and

acquired by a sCMOS camera (Hamamatsu,

C11440-22CU) with a 50 ms exposure time.

Next, merocyanine 540 (MC540, Invitrogen)

was introduced to cell media to image the cell

membranes via PAINT (Point Accumulation

for Imaging in Nanoscale Topography) [32].

Similar to AF488-PD, localizations were only

acquired when MC540 temporarily docked

with the cell membranes and emitted bright

fluorescence, thereby enabling the co-

localization of prodrug molecules relative to

cell membrane with nanoscale resolution.

MC540 was excited with a 561-nm laser (Sapphire, Coherent) at a peak intensity of 2.7

kW∕cm2 at the sample, and images were

acquired with a 50 ms exposure time. The

single AF488-PD and MC540 molecules were

identified and localized by two-dimensional

Gaussian fitting using an ImageJ plug-in

ThunderSTORM [33].

The super-resolution images of prodrug and

cell membranes consist of the sum of all

localizations from AF488-PD and MC540

respectively, binned as a 2D histogram with bin

size 58.5×58.5 nm2. AF488-PD localizations

over time were grouped into a trajectory when

present in consecutive frames (up to a 3-frame

gap) within a distance of 0.59 μm. The

trajectories were plotted using custom software

(MATLAB, MathWorks). No trajectory

analysis was performed for the endothelial

filopodia.

Results

Spectroscopic characterization of synthesized prodrugs

Absorption and fluorescence spectra and

decay characteristics of the materials were

measured in a water solution. DOX and DOX-PD had similar absorption spectra (Figure 1A).

The spectra for αvβ3-DOX-PD PFOB NP was

obscured by light scattering.





Figure 1. Spectroscopic characterization of doxorubicin (DOX), doxorubicin prodrug (DOX PD), and αvβ3-integrin

targeted prodrug nanoparticles (αvβ3-DOX-PD PFOB-NPs) in water: Absorption spectra of the materials (A). The absorption

of doxorubicin prodrug in nanoparticle formulation is obscured by light scattering. Presence of doxorubicin is confirmed in

the corresponding fluorescence emission (B). Fluorescence decay characteristics of the materials (C). DOX decay can fit to

mono-exponential decay of 1.09 ± 0.02 ns. DOX-PD can fit to bi-exponential decay with τa = 1.21 ns (47%); τb = 0.69 ns

(53%). DOX-PD NP can fit to bi-exponential decay with τa = 3.78 ns (22%); τb = 1.34 ns (78%). Data represent mean ± sd,

n=3 measurements. IRF – Instrument response function.

All drug formulations showed similar

fluorescence emission spectra, which

corroborated the presence of doxorubicin in the

nanoparticles (Figure 1B). The emission peak at

~556 nm for doxorubicin in the αvβ3-DOX

PFOB NP was relatively more prominent. This

observation may reflect the unique interactions

of the doxorubicin molecules with the core

PFOB molecules that penetrate to the particle-

water interface [34].

As shown in Figure 1C, free DOX

fluorescence fit a mono-exponential decay of

1.09 ± 0.02 ns (χ2 = 1.06), consistent with

literature findings. DOX-PD decay best fit a bi-

exponential decay model with nearly equal

contributions from a fast lifetime component τa

= 0.69 ± 0.06 ns (53%) and a slow lifetime

component of τb = 1.21 ± 0.03 ns (47%),

indicative of a heterogeneous system (χ2 =

1.007). The amplitude average lifetime (τamp) was calculated to be 0.96 ± 0.04 ns. Dynamic

light scattering analysis of a DOX-PD in water

revealed a mixed suspension of free prodrug

molecules and spontaneously formed prodrug

micelle aggregates (~1600 nm). Fluorescence

lifetime of doxorubicin bound to biological

molecules (lipid membranes, DNA) and while

encapsulated in nanoparticles increases as the

molecules are shielded from external

fluorescence quenchers and by - stacking

[35-38]. Hence, the slower lifetime component

from DOX-PD suspension was attributed to the

micellar fraction of the drug distribution where

the anthracycline portion of DOX-PD was

shielded from surrounding water molecules.

The faster lifetime of 0.69 ns can be attributed

to the free DOX-PD molecules in water. This

value is slightly different than that of free DOX

in water, reflecting the effect of drug

conjugation with the lipid molecule.

Fluorescence decay of αvβ3-DOX-PD PFOB

NP fit a double exponential decay model with





components τa = 3.78 ± 0.03 ns (22%) and τb =

1.34 ns (78%) (χ2 = 1.03). The majority of the

αvβ3-DOX-PD PFOB NP had the faster

fluorescence lifetime component (1.34 ns)

similar to that of DOX-PD in the micellar

fraction (1.21), indicating similar hydrophobic

interactions. The slower lifetime fractional

component (3.78 ns) is likely related to

hydrophobic interactions of DOX-PD within

the nanoparticle lipid surfactant intercalated

between PFOB molecules.

Cellular uptake and cytotoxicity of αvβ3-DOX-PD PFOB NP

The cellular distribution of DOX presented as

the free drug and delivered within nanoparticle

surfactant in prodrug form was observed with

fluorescence microscopy (Figure 2).

Figure 2. Cellular internalization and cytotoxic effects of αvβ3-DOX-PD PFOB-NP: Internalization of DOX and αvβ3-

DOX-PD PFOB-NP in 2F2B cells incubated with reagents for 30 min. Red shows doxorubicin fluorescence signal and blue

is nuclear counterstain, overlay is purple. (A) 2F2B cells treated with free DOX showing most red signal localized into nucleus

(white arrow). (B) 2F2B cells treated with αvβ3-DOX-PD PFOB-NP showing signal diffuse through cytosol, membrane

(yellow arrow) and nucleus (white arrow). (C) MTT cytotoxicity assay after 48 h of treatment, showing αvβ3-DOX-PD PFOB-

NP was significantly more cytotoxic than the equivalent dose of free DOX (p<0.05) while drug-free NP had no effect on cell

proliferation. Statistical analysis was performed using unpaired Student's T test and p values were assigned.

Benign 2F2B mouse endothelial cells were

exposed to DOX and αvβ3-DOX-PD PFOB NP.

Free DOX internalized and localized rapidly to

cell nuclei within 30 min of treatment with

negligible cytosolic retention noted (Figure

2A). DOX-PD delivered by αvβ3-targeted NP

bound to the cell membrane periphery, entered

the cell via the CFDD mechanism and rapidly

dispersed into the cytosol membranes. Lipase-

released DOX was observed in the nucleus (Figure 2B). Circumventing cell internalization

with endosome sequestration and escape, a

common pathway for nanoparticle drug

delivery, DOX-PD dispersed throughout the

cytoplasmic membrane. Enzymatically

liberated DOX accumulated in the nucleus.

The cytotoxic bioactivity of αvβ3-targeted

DOX-PD PFOB NP referenced to equimolar

free DOX was assessed in 2F2B mouse

endothelial cells stimulated with angiotensin II

using the MTT cytotoxicity assay. The control

αvβ3-no-drug NP had no impact on cell

proliferation (Figure 2C). Both free DOX

added to the media and αvβ3-DOX-PD PFOB

NP were cytotoxic (Figure 2C). At 48 h αvβ3-

DOX-PD PFOB NP decreased cell proliferation

by 44% (P<0.01) as compared to the control

αvβ3-no-drug NP. An equivalent dose of free

DOX decreased cell proliferation by 19%

(P<0.05) compared to the control. The αvβ3-

DOX-PD PFOB NP was significantly more

toxic than equivalent amounts of free DOX (P<0.05). These data showed that the Sn-2

coupling of DOX to the phosphatidylcholine

lipid backbone did not compromise drug net

bioactivity and suggested that continued

liberation of DOX from the DOX-PD reservoir

retained in the cytoplasm over 48 hours

contributed to effectiveness.

Temperature dependent cellular uptake of αvβ3-DOX-PD PFOB NP

Fluorescence imaging of prodrug trafficking

performed in normal proliferating endothelial

cells was reflective of previous anti-





angiogenesis experiments [10, 12, 39, 40] with

Sn-2 prodrugs. As opposed to normal

proliferating endothelial cells, the next

experiments explored Sn-2 prodrug behavior in

abnormal human C-32 melanoma.

Drug uptake from αvβ3-DOX-PD PFOB NP

was studied in cultured C-32 cells at 37 °C and

at 4 °C, the later to induce cold suppression of

ATP-dependent mechanisms (Figure 3).

Figure 3. Effect of temperature on cellular uptake of αvβ3-DOX-PD PFOB-NP: Comparison of cellular uptake of αvβ3-

DOX-PD PFOB-NP after 1 h incubation at 37 degrees (A) and 4 degrees C (B). Control untreated cells show near zero signal

(C). Quantified background corrected whole cell fluorescence (mean ± sem, n=5 cells for each of the 3 independent

experiments) showing internalization is reduced at 4 degrees C (D).

DOX-PD transfer from the bound

nanoparticle surfactant to the outer cell

membrane leaflets was similar under both

temperature regimens. While transfer of DOX-

PD from the outer to inner cell membrane

leaflets was rapid and unimpaired at 37 °C, at 4

°C DOX-PD delivery to the inner leaflet and

contiguous intracellular membranes was

reduced but notably present well-above

background fluorescence. This was in

contradistinction to previous studies using

rhodamine-PE as a drug surrogate in which

translocation of the fluorescent lipid from the

outer to inner cell membrane leaflets was

blocked at 4 °C. These data suggest that DOX-

PD and other PC-PD will translocate between

membrane leaflets by ATP-dependent and

ATP-independent mechanisms and slowly

diffuse into cells in the absence of energy.

Pixel-by-pixel fit of the raw decay data shows

gradient of amplitude average lifetime τamp

throughout the cells (Figure 4B). Untreated

cells showed minimal background signal under

these imaging conditions (Figure S4). We

performed cell compartment specific analysis

of our lifetime data as below (Figure S5).

Nucleus

Nuclei of cells treated with DOX and αvβ3-

DOX-PD PFOB-NPs showed similar lifetime

distribution suggesting that DOX liberated

from lipid prodrug retained the same dsDNA

binding character as the native compound

(Figure S3). Previous studies have reported

DOX bound to dsDNA exhibits higher lifetime

(2.2 ns – 1.3 ns, [37, 38, 41]). Upon fitting

decays from nuclear ROIs using bi-exponential

model, a major component of fast lifetime 1.44

ns ± 0.06 ns (80% ± 4%, n=52) was detected

reflecting dsDNA-bound DOX. A minor slow

lifetime component (3.80 ± 0.24 ns) in the

nucleus was attributed to noise from

backscattering of excitation light and auto-

fluorescence, which was also noted for

untreated cells (Figure S4). τamp in the nuclei

decreased steadily between 1 h (2.11 ± 0.04 ns)

and 8 h (1.82 ± 0.02) post-incubation (Figure

4C, P<0.001). This observation was previously

observed by others and attributed to drug-

induced apoptosis with consequent remodeling

of chromatin network in the nucleus [41].





Figure 4. Fluorescence lifetime imaging of C-32 cells treated with αvβ3-DOX-PD PFOB-NP. Fluorescence intensity images

of C-32 cells with αvβ3-DOX-PD PFOB-NP for 1, 4, 8, and 24 h, showing doxorubicin fluorescence from cell cytoplasm and

nucleus at all time points. A perinuclear space with high signal intensity is observed in all cells (red arrows) indicating

accumulation of drug (A). Corresponding fluorescence lifetime images generated by pixel-by-pixel fitting of decay data show

a gradient of τamp in different cell compartments (B). Quantification of τamp from nuclear compartments shows time dependent

decrease (n= 7-18 cell nuclei at each time point, 3 independent experiments, p<0.001) suggested drug induced apoptosis (C).

Percentage contributions of slow and fast components in the bright perinuclear region (τamp = 1.8 ± 0.2 ns) compared to the

rest of the cytosol ((τamp = 2.9 ± 0.3 ns)) for cells treated with αvβ3-DOX-PD PFOB-NP for 8 h (D). Bright perinuclear ROIs

show greater contribution of fast lifetime than cytosolic regions outside it (mean ± sem, n= 15 cell ROI measurements).

Statistical analysis was performed using unpaired Student's T test and statistical significance was assigned for p<0.001,

represented by ***.

Cytoplasm

The cytoplasm of cells treated with αvβ3-

DOX-PD PFOB-NPs demonstrated fluores-

cence signal at all time points with consistently

high signal noted in the perinuclear space

(Figure 4A, red arrows). This region coincides

with the location of Golgi/ER, which is

associated with enriched levels of cytosolic

phospholipase A2 (cPLA2) activity [42].

Comparisons within and outside the bright

perinuclear regions showed lower τamp (1.8 ns

± 0.2 ns, n=15 ROI) in the perinuclear regions

than the later (τamp = 2.9 ns ± 0.3 ns, n=15 ROI).

The lower τamp observed in bright perinuclear

region was due to greater contribution from a

faster lifetime component (1.1 ns, 82% ± 3%)

suggesting an abundance of free DOX

molecules. In the surrounding cytosol, this

faster component was much less in proportion

(1.06 ns, 52% ± 8%), indicating lower amounts

of free DOX (Figure 4D). The slower lifetime

components from both perinuclear and

surrounding regions was attributed to a

combination of background (as observed in cell

nuclei) and from bound drug (as observed in

literature and our spectroscopic studies). By

extension, the collective data suggested that

phospholipase release of Sn-2-linked

doxorubicin from the PC backbone was more

concentrated in the perinuclear region than in

the general cytosol.





Super-resolution microscopy of cellular binding and internalization of prodrug molecules

Single-molecule super-resolution microscopy

(SRM) captures images of fluorescent

molecules with nanometer resolution beyond

the optical diffraction limit. The AF488-PD,

which contains fluorescent AF488 molecules

conjugated to the phospholipid at the Sn-2

position, preferentially intercalated into the

outer membrane leaflet of cells.

Figure 5. Early-stage binding of AF488-PD on the membrane and internalization in 2F2B cells. (A) Bright-field and super-

resolution images of AF488-PD (red, after 5 min incubation) and cell membrane probe MC540 (green) on the membrane of

a 2F2B cell. The overlay images are focused at different heights above the coverslip. The arrows indicate the colocalized

active binding sites of AF488-PD and MC540 on the membrane. (B) Both transient binding and diffusion (trajectories on the

right part of the image) of single AF488-PD molecules were observed on the cell membrane after 5 min incubation.

Trajectories on the filopodia were not shown. Magnified views of the boxed regions shown in insets: (i) scatter plot of single

AF488-PD molecules (red) binding to filopodia, (ii, iii) representative trajectories of single AF488-PD molecule diffusing on

the cell membrane with the total length of 3.50 s and 1.65 s, respectively; color-coded for time. (C) Fluorescence images of

AF488-PD (red) after a 12-hour incubation at different heights above the coverslip, showing intracellular accumulation of

the prodrug. PAINT images of MC540 (green) indicate the contour of the cell membrane. CS, coverslip; MEM, membrane;

CY, cytoplasm; NU, nucleus.





In 2F2B endothelial cells, AF488-PD (Figure

5A, red) was imaged within the cell membrane

after 5 min of incubation, co-localizing with

another membrane marker dye, MC540 (Figure

5A, green). As noted in the bright field images

shown in Figure 5A, SRM of AF488-PD and

MC540 both accumulated in the outer cell

membrane at different z heights (1, 2, 3 μm)

above the coverslip. These merged PAINT

images suggested that the PC prodrug

inherently incorporated into the cell membrane

with little spatial restriction between the base of

the cell on the coverslip and the apical surface

fully exposed to the bulk media. In addition to

the primary cell body surface membrane,

AF488-PD also incorporated into 2F2B cell filopodia processes as noted in the magnified

image of the box (i) in Figure 5B. In

contradistinction, standard epifluorescence

microscopy imaging showed that the PE-

rhodamine did not broadly intercalate into the

outer phospholipid cell membrane but was

primarily associated with the cell along the

coverslip region where cell adherent molecules

are highly expressed and engaged (Figure S6).

This observation was consistent with the report

of Partlow et al, which associated PE-

rhodamine uptake with specific lipid-raft

membrane regions [4].

The diffusion of single molecules of AF488-

PD through the cytosol was imaged using

single-molecule fluorescence imaging

(Supplementary video S1). The trajectories

shown in Figure 5B illustrated the trackable

paths of single prodrug molecules prior to

photobleaching, diffusion beyond the focal

plane of the microscope, or enzymatic dye

liberation into the cytosol. Two representative

trajectories are presented in the magnified

images, box (ii) and (iii), in Figure 5B.

Individual freely-diffusing prodrug molecules

were also observed at the coverslip level of the

cell, which likely reflected the rapid mobility of

cell membrane phospholipid constituents [43].

With longer incubation times at 37 °C more

AF488-PD entered the membrane from the

media and internalized into the cell. Figure 5C

shows the fluorescence images of 2F2B cells

after incubation with AF488-PD for 12 h. At

this stage, AF488 fluorophores accumulated

broadly within the internal cell membranes

particularly in the perinuclear region,

corroborating the fluorescence microscopy

observation. Fluorescence images obtained at

different z heights (Figure 5C at 0.5, 2.5, 6.5 μm

above the coverslip) showed dye throughout the

cell with the intensity of the light signal

proportional to the proximity of AF488-PD to

the microscope’s focal plane, while the

autofluorescence background from 2F2B cells

was low (Figure S7). Diffuse red signal

associated with AF488-PD was noted beyond

the microscope’s depth of field. The rapid

motion of freely diffusing liberated AF488

molecules were not resolved in the cytoplasm

(Figure 5C).

Discussion

In vivo efficacy of Sn-2 PC prodrugs

incorporated into the surfactant of lipid-

encapsulated nanoparticles has been

demonstrated previously [9-12]. These studies

showed that the Sn-2 PC PD motif inhibited

significant premature drug loss and drug

metabolism during circulation to the targeted

pathology. However, evidence corroborating

prodrug avoidance of internalization and

sequestration within the endosome/lysosome

pathway versus CFDD-mediated translocation

into intracellular membrane distribution with

subsequent enzymatic drug liberation was

lacking. In the present series of experiments,

PC prodrug delivery into cells and intracellular

trafficking was investigated using three

complementary optical imaging methods:

classic fluorescence microscopy, fluorescence

lifetime imaging microscopy, and single-

molecule super-resolution microscopy. The

overarching result suggests that the PC

prodrugs act as mimics of phosphatidylcholine

with regard to cell membrane uptake,

membrane leaflet translocation, intracellular

distribution, and enzymatic metabolism in

normal and cancerous cell lines.

In prior research with rhodamine-PE as a lipid

drug surrogate in αvβ3-targeted lipid-

encapsulated NP, access into cells was confined

to lipid rafts and transfer of the fluorescent PE

from the outer to inner cell membrane leaflets

was dependent on ATP, consistent with the a

well-known amino-phospholipid flippase

transport protein mechanism. In ATP-starved

situations, such as hypoxic cancer cells,

targeted delivery of PE-based prodrugs could



https://precisionnanomedicine.com/article/15



be compromised. In contradistinction, PC-

prodrugs intercalated throughout the outer cell

membrane leaflet, including filopodia

processes and translocated to the inner leaflet

rapidly at 37 C. ATP-suppressing cold

treatment (4 C) partially inhibited the outer to

inner membrane leaflet translocation of PC-

prodrug, but substantial drug transferred into

the inner leaflet and then migrated throughout

the cytosolic membranes. These data suggest

that DOX-PD and other PC-PD will translocate

between membrane leaflets by ATP-dependent

and ATP-independent mechanisms and slowly

diffuse into cells in the absence of energy.

Moreover, spatially unrestricted cell entry

combined with ATP independent translocation

into the cell are both favorable for drug

delivery.

PC prodrugs broadly disseminated into

cytosolic membranes overtime, but a

significant intracellular trajectory was noted in

the lipid membranes of vesicles, likely

endosomes, trafficking toward the perinuclear

nuclear Golgi/ER region, a pathway well-

documented for native PC [26]. The

perinuclear Golgi/ER region has enriched

concentrations of cytoplasmic phospholipase

A2, a lipase involved in the degradation and

recycling of phosphatidylcholine [42, 44].

Enhanced local perinuclear enzymatic

liberation was demonstrated with doxorubicin

Sn-2 prodrug (DOX-PD) and fluorescence

lifetime microscopy. In perinuclear Golgi/ER

region, increased concentrations of free

doxorubicin were estimated. In other

cytoplasmic membrane regions with prevalent

DOX-PD retention, substantially lower

concentrations of released drug were estimated.

Fluorescence lifetime imaging further

demonstrated that the PC-released doxorubicin

entered the nucleus and bound to dsDNA

similar to native doxorubicin. In culture, free

doxorubicin entered cells rapidly and localized

to the nucleus with negligible cytoplasm

retention. By comparison, DOX-PD entered

the cell after NP targeting and a visually

significant portion of the drug rapidly bound

within the nucleus. However, unlike free DOX,

less drug was observed in the nucleus and a

significant proportion of the prodrug remained

within the cytoplasmic membranes.

Nevertheless, the biopotency of the native and

prodrugs were at least equivalent with regard to

cell proliferation when measured by MTT assay

at 48 hours. If the drug cytotoxicity is directly

correlated to the compound bound to dsDNA,

then the residual prodrug within the

intracellular membrane may have constituted a

slower release depot with continued drug liberated over the 48-hour experiment

increasing nuclear DOX delivery.

Alternatively, the slower translocation of DOX-

PD to the nucleus with residual cytoplasm

retention may have increased the

antiproliferative contributions of other

pathways of doxorubicin toxicity, such as in the

mitochondria [45].

In the present study, definitive confirmation

of PC-PD migration to the Golgi, specifically,

was not established and tracking to the

endoplasmic reticulum or both organelles in the

perinuclear region could not be differentiated.

Additionally, the potential role of doxorubicin

exocytosis when delivered as the prodrug

versus free drug form was not studied.

Attention was placed on the binding of liberated

DOX to DNA over time, which evolved slowly

from the retained cytosol membrane depot.

Finally, the MTT assay was used to assess cell

apoptosis. Although careful washing of residual

doxorubicin prior to the addition of assay

reagents to the cell culture was performed, a

complementary caspase assay might additionally inform whether the treated cells

cycled into apoptosis versus the control.

Conclusion

Sn-2 lipase labile phospholipid prodrugs in combination with the contact-facilitated drug

delivery mechanism have been demonstrated for targeted nanoparticle therapy in vivo. In this report,

traditional fluorescence microscopy, fluorescence lifetime imaging, and single-molecule super-

resolution imaging techniques were used to elucidate the intracellular trafficking and metabolism details

of PC prodrugs for the first time. PC prodrugs incorporated broadly into the outer cell membrane leaflet

over the cell body and filopodia, and translocated to the inner membrane leaflet by ATP-dependent and ATP-independent mechanisms. Nanoparticles were not sequestered into endosomal vesicles, but the PC





prodrugs delivered into the cells populated vesicle membranes that trafficked to the perinuclear

Golgi/ER region as well as distributed through the intracellular membranes in the cytosol. As an

example, doxorubicin-prodrug tracked to the Golgi/ER region where the highest level of liberated drug

was measured compared to the general cytosol. The released anthracycline displayed similar

fluorescence lifetimes when bound to dsDNA as the native. In contradistinction to free doxorubicin,

which transferred directly to the nucleus after cell uptake, a significant portion of DOX-PD remained

within cytosolic membranes. Nevertheless, the biopotency of the native and prodrugs were at least

equivalent with regard to cell proliferation at 48 h, which suggests that the retained DOX-PD may have

constituted a prolonged release reservoir that enhanced effectiveness over two days. While the specific

fluorescence lifetimes measured in the present study have limited relationship with values for in vivo

imaging, this study provided the first evidence-based understanding of intracellular Sn-2 PC prodrug

fate.

Acknowledgements

Partial funding support was received from the Barnes-Jewish Hospital Research Foundation, St Louis,

MO, 63110 and from the NIH: HL122471, HHSN-268201400042C, HL112518, HL112518,

HL113392, CA199092, CA154737, S10 OD016237, P50CA094056. DM was supported by the

Imaging Sciences Pathway graduate student fellowship at Washington University in St. Louis under

grant number NIH T32 EB014855.

Disclosure:

Washington University (GL, DP) has intellectual property rights related to the prodrug technology

described in this manuscript. The remaining authors have nothing to disclose.

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Quote as: Maji D, Lu J, Sarder P, Schmieder AH, Cui G, Yang X, Pan D, Lew MD, Achilefu S, Lanza

GM. Cellular Trafficking of Sn-2 Phosphatidylcholine Prodrugs Studied with Fluorescence Lifetime

Imaging and Super-resolution Microscopy. Prec. Nanomed. 2018 July;1(2):127-145

DOI:10.29016/180730.1

Video S1. Video clip showing the single-molecule fluorescence images of AF488-PD molecules on a

2F2B cell within the same field of view as in Figure 5B in the main text. The trajectory from 0.55 to

4.00 s indicates the diffusive path of a single AF488-PD molecule on the cell membrane (see also,

Figure 5B, inset (ii) in the main text).




https://precisionnanomedicine.com/static/video/Supplementary%20Video%20S1%20prnano_132018.mp4

SUPPLEMENTARY INFORMATION

Figure S1. Chemical structures and characterization for Doxorubicin PD, AF488-PD and PE-Rhodamine. (A). Chemical structure of doxorubicin prodrug as described in methods. Chemical Formula: C60H91N2O20P, ESI-MS M+H+ Cal. 1190.63, exp.1191.6 (Shimadzu LCMS 2010A, Kyoto, Japan). (B). Chemical structure of AlexaFluor-488-prodrug (AF488-PD) as described in methods. The structure produced in high purity because the purportedly high purity starting materials, AF-488 dye and Paz-PC, were both unexpectedly impure as purchased and received. The PAz-PC, synthesized by NOF America from the lysophosphatidylcholine backbone and the addition of nonanedioic acid at Sn-2 contained significant free nonanedioic acid. AlexaFluor-488 cadaverine purchased from Thermo Fisher Scientific presented two N-(5-aminopentyl)formamide arms instead of one on the benzyl ring. As a consequence, the free nonanedioic acid also coupled to the extra N-(5-aminopentyl)formamide to create a second hydrophobic anchor. Synthesis of this compound into lipid membranes was unaffected and it was used as surrogate biomarker for single-molecule tracking. The final chemical formula was C74H113N7NaO23PS2+2H+, ESI-MS M+H+ Cal. 793, exp.793.85 (Shimadzu LCMS 2010A, Kyoto, Japan). (C). Chemical structure of Rhodamine-phosphatidylethanolamine (PE-Rhodamine, Avanti Polar Lipids). It was previously used by Partlow et al. [1].

1

Figure S2. Schematic of the imaging system used for single-molecule super-resolution fluorescence imaging. Only one channel of the polarized images was captured and used. BP1-3, bandpass filters; QWP1-2, quarter wave plates; M1-7, mirrors; DM1-2, dichroic mirrors; L1-5, lenses; KL, wide-field lenses; OL, objective lens; TL, tube lenses; PM, pyramidal mirror; PBS, polarizing beam splitter; SLM, spatial light modulator.

Figure S3. Images of C-32 melonoma cells treated with DOX and αvβ3-DOX-PD PFOB-NPs for 1 h. DOX treated cells (A: intensity (top), fast lifetime (bottom)) exhibit strong nuclear signal while the later showed strong fluorescence throughput (B: intensity (top), fast lifetime (bottom)), and the untreated cells showed minimal background signal (C, intensity). Cell nuclei in both groups showed similar fast lifetime distribution as shown by the color-coded images and lifetime histogram (D), suggesting that DOX liberated from lipid prodrug retained the same dsDNA binding character as the native compound.

Figure S4. Under the imaging conditions, untreated C-32 cells gave minimal background signal due to cell auto fluorescence and backscattering of excitation laser (A). This signal could be visualized by changing the intensity scale from 0-2000 to 0-200 events (B), and contributed <10% of real fluorescence signal under these conditions. Lifetime of this signal was broadly spread with an average of around ~ 3.5 ns as shown in the histogram (C, D).

Figure S5. Representative fluorescence lifetime image analysis of C-32 cells treated with αvβ3-DOX-PD PFOB-NPs as shown in Figure 4 of main text. The average lifetime image obtained after pixel-by-pixel fit of decay using a bi-exponential decay model (A). The corresponding image displaying the chi-square values obtained shows goodness of fit (B). Nuclear region of interest (C, left) and corresponding fluorescence decay with its bi-exponential fit (C, right, χ2=1.04). Similar analysis is shown for a region of interest corresponding to the bright perinuclear space, χ2 =1.006. (D, left and right)

Figure S6. Epifluorescence images of a 2F2B cell at different z heights (0.5, 2.0, 5.0 µm) above the coverslip after incubation with 1.4 µM PE-rhodamine at 37 ºC for 1 hour.

Figure S7. Epifluorescence images of (A) autofluorescence from an untreated 2F2B cell and (B) a 2F2B cell after incubation with 4.0 µg/mL AF488-PD at 37 ºC for 12 hours.

Video S1. Video clip showing the single-molecule fluorescence images of AF488-PD molecules on 2F2B cell within the same field of view as in Figure 5B in the main text. The trajectory from 0.55 to 4.00 s indicates the diffusion path of single AF488-PD molecule on the cell membrane (see also, Figure 5B, inset (ii) in the main text). SUPPLEMENTAL INFORMATION BIBLIOGRAPY 1. Partlow KC, Lanza GM, Wickline SA. Exploiting lipid raft transport with membrane targeted nanoparticles: a strategy for cytosolic drug delivery. Biomaterials. 2008; 29: 3367-75.

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Cellular Trafficking of Sn-2 Phosphatidylcholine Prodrugs